The authors would like to thank the Calvin Kuo Laboratory at Stanford University for providing HEK293 cells stably transfected with either HA-mouse Noggin-Fc or HA-mouse Rspo1-Fc. We would also like to thank Dr. Dean Tang for allowing us access the fluorescent dissection microscope in his laboratory. This work was supported by CA179907 to D.W.G. from the National Cancer Institute. Shared resources at Roswell Park Comprehensive Cancer Center were supported by National Institutes of Health Cancer Center Support Grant CA016056.

Animal procedures described here were performed with the approval of the Institutional Animal Care and Use Committee (IACUC) at the Department of Laboratory Animal Resources, Roswell Park Comprehensive Cancer Center, Buffalo, New York.
NOTE: Male mice to be dissected to isolate prostates or prostate tumors for generation of organoids should have at least reached the age of sexual maturity &#8212; about 8-10 weeks of age. Specific ages of mice can vary amongst studies. Some factors to consider when choosing age include age-dependent changes in prostate cell populations, age-dependent expression of specific promoter-driven Cre transgenes, and rate of prostate tumor progression in a particular GEMM.
1. Dissection and Imaging of Mouse ProstateTumor and Metastatic Tumors
<ol>
	<li>Preparations
	<ol>
		<li>Obtain necessary sterile dissection tools. Stage dissection area on clean sterile surface with a 15 cm ruler, precision balance, analytical balance, spray bottle with 70% ethanol, phospho-buffered saline (PBS), and paper towels.</li>
		<li>Preparation of fixative solutions for visceral organs and bones not to be used for organoid generation
		<ol>
			<li>For visceral organs: Prepare 4% paraformaldehyde (PFA) in PBS. For one mouse, make 20 mL of 4% PFA, aliquot into two 15 mL conical tubes and keep at room temperature until use.</li>
			<li>For long bones: Aliquot 10 mL of pre-made 10% neutral buffered formalin (NBF) in one 15 mL conical tube and keep at room temperature until use.</li>
		</ol>
		</li>
		<li>Obtain untreated 10 cm dishes and fill with non-sterile PBS &#8212; these will serve as temporary containers for organs during fine dissection using a dissection microscope. 
		NOTE: Sterile tools are used for dissecting tissues for generating organoids. Non-sterile tools are used for the initial incision and dissection of the hind limbs.</li>
	</ol>
	</li>
	<li>Euthanasia and initial incision
	<ol>
		<li>Euthanize the mouse by CO2 asphyxiation using a 2.0 L/min flow rate for 5 min. Remove the mouse from the cage and perform cervical dislocation. Use the precision balance to measure the mouse&#39;s body weight and record.</li>
		<li>Place the mouse on top of a paper towel and orient on the dissection surface ventral side up with the mouse&#39;s head facing away from the investigator. Affix the mouse to the board by stretching out its limbs and piercing each of the forepaws and hind paws with one disposable needle.</li>
		<li>Using a spray bottle, douse the mouse&#39;s fur with 70% ethanol. Using non-sterile dissection scissors and straight forceps, pinch just above the mouse&#39;s penis and make a small incision through the fur only.</li>
		<li>Continue the incision midline up to the mouse&#8217;s neck through the fur only. Make bilateral incisions from the point of the initial incision through the mouse&#39;s ventral plane through the fur only.</li>
		<li>Grasp the fur and carefully pull it away from the skin of the mouse. Pin down the fur to the board to allow access to both the abdominal and thoracic cavities.</li>
	</ol>
	</li>
	<li>Extraction of male urogenital system en bloc
	<ol>
		<li>Using sterile dissection scissors and straight forceps, carefully cut through the skin about 0.75 cm above the rectum. Continue the incision midline up to the ribcage without disturbing any organs in the abdominal cavity. Carefully pull the skin away and pin to the board to expose the entire abdominal cavity. 
		NOTE: Do not allow these instruments to touch the outside of the mouse or any surface in the staging area, as these tissues will be used to generate organoids.</li>
		<li>Dissect the urogenital system and other organs according to the diagram in Figure 1A, with numbers indicating the order the organs will be removed from the mouse.</li>
		<li>Find the bladder, then grasp the fat pad to either the left or right and pull upward to expose the testicle. Carefully dissect away the testicle from the rest of the urogenital region and put aside. Do the same procedure on the other side. 
		NOTE: Achieving the optimal tissue quality and detailed tumor characterization of prostate tumors requires that the entire urogenital region be removed from the mouse en bloc19. It is recommended that the urogenital region be removed first (Figure 1A).</li>
		<li>Grasp the bladder and carefully pull up so that the urogenital region lifts together, exposing the urethra underneath. While holding the bladder, orient the scissors so they are against the underside of the dorsal prostate and cut the urethra. The entire urogenital region will then release from the abdominal cavity.</li>
		<li>Put the urogenital region in a 10 cm dish filled with PBS. If still filled with urine, drain the bladder by making a small incision. Using the analytical balance, weigh the urogenital system and record.</li>
	</ol>
	</li>
	<li>Extraction of pelvic lymph nodes, spleen, liver, kidney, lung, tibia, and femur
	<ol>
		<li>Removing the urogenital system exposes the pelvic lymph nodes, positioned right behind the urogenital system (Figure 1A) and on either side of the spine. The lymph nodes will only be visible if they contain metastatic lesions or if there is local inflammation. Orient the straight forceps underneath the lymph node and pull up to remove the lymph node. Do the same procedure on the other side.</li>
		<li>Grasp the rectum with the straight forceps and cut. Pull on the rectum to unravel the entire colon and small intestine, looking for metastatic lesions in the mesenteric lymph nodes. When the entire ileum is removed, continue to pull on the duodenum to expose the stomach. Cut the esophagus to completely remove the stomach and discard. If any metastatic lesions in the lymph nodes are observed, carefully dissect away from the intestine and store in a 10 cm dish of PBS.</li>
		<li>Exposing and removing the stomach will pull the spleen from the dorsal side of the abdomen (Figure 1A). Remove the spleen and place in 4% PFA. The spleen serves as a staining control for hemotoxylin as it is highly cellular. 
		NOTE: Visceral tissues not to be used for organoid generation will be fixed overnight in 4% PFA, washed with PBS, and then placed in 70 % ethanol.</li>
		<li>Remove the liver in the upper part of the abdomen (Figure 1A). Based on the metastatic load in the liver, individual lobes can be dissected, or the entire liver can be removed by carefully cutting along the diaphragm. (Do not cut through the diaphragm at this time). Place the liver in a 10 cm dish of PBS.</li>
		<li>Removing the liver will fully expose the kidneys on either side of the spine (Figure 1A). Remove the kidneys &#8212; along with the renal lymph nodes, if the nodes have metastatic lesions &#8212; by placing the straight forceps underneath the kidney and pulling up. Do the same procedure for the other side.</li>
		<li>To expose the thoracic cavity, carefully cut the diaphragm along the ribcage. Piercing the diaphragm will release the negative pressure in the thoracic cavity and expose the heart and lungs (Figure 1A).</li>
		<li>Grasp the sternum and pull up to open the thoracic cavity further. Observe the ventral face of the thoracic cavity along the rib cage for metastatic lesions in the thoracic lymph nodes and dissect, if present.</li>
		<li>While still holding the sternum, cut away the ventral rib cage to access the heart and lungs. Grasp the heart, pull up and cut underneath the lungs. To fully remove the heart and lungs en bloc, cut all anterior blood vessels and the trachea. Place the tissue in PBS and carefully remove the heart without damaging the lung tissue or lung metastatic lesions.</li>
		<li>Take the non-sterile straight forceps and dissecting scissors and cut the hind leg at the head of the femur. Carefully grasp the femur and remove the hind leg from the fur.</li>
		<li>Place the hind leg on a paper towel and grasp the hind paw. Use the single edge razor blade to scrap/cut away all muscle and connective tissue from the tibia and femur. Remove the femur from the tibia by cutting posterior to the patella and remove the tibia by removing the hind paw. Place the tibia and femur in 10% NBF. Do the same procedure on the other side. 
		NOTE: For the purposes of examining the long bones of the mouse for metastatic lesions, the fibula does not need to be intact. The long bones will be fixed in 10% NBF for one week, then decalcified using neutral EDTA solution20. After three weeks, the bones will be transferred to 70% ethanol.</li>
		<li>After removing the hind limbs, take an ear or tail cutting for future genotyping. Discard the mouse carcass and all tissue not to be fixed or used for organoid generation. &#160;</li>
	</ol>
	</li>
	<li>Dissection of prostate tumor 
	NOTE: Dissection of mouse prostate lobes can only be achieved using a dissecting microscope19. However, the prostate tumor load can be so high that individual lobes cannot be distinguished, and dissection can be carried out without a microscope. Nevertheless, the full protocol for dissection of individual prostate lobes is described below.
	<ol>
		<li>Place the urogenital region in a 10 cm dish of PBS under a dissecting microscope and orient the ventral side up with the bladder and seminal vesicles facing away from the investigator. All manipulation of the urogenital region should be done using sterile instruments.</li>
		<li>Assess the general phenotype of the prostate tumor-&#160;most likely, either a fluid-filled or solid tumor will be observed in PCa GEMMs (Figure 2A, B). 
		NOTE: Fluid-filled tumors are often located in the anterior prostate region (Figure 2A). Fluid filled tumors are made up of primarily connective tissue with scant epithelial components &#8212;&#160;thus these tumors are not optimal for organoid generation because of the low number of tumor epithelial cells, as will be discussed in the representative results section.</li>
		<li>If fluid filled tumors are present, poke a small hole in the tumor. Place the urogenital region in a new 10 cm dish of PBS.</li>
		<li>Take a pair of straight forceps in the non-dominant hand and curved forceps in the dominant hand. Flip the urogenital region to its dorsal face. Look for the proximal prostate region (Figure 1B), which can be identified by the pink/red color of the urethra. Grasp the urethra and hold firmly so to manipulate the urogenital tissue with the curved forceps. 
		NOTE: During prostate dissection, always take note of the bladder&#39;s location, as this is the simplest way to locate the individual prostate lobes (Figure 1B).</li>
	</ol>
	</li>
	<li>Removal of non-prostate tissue from the urogenital region
	<ol>
		<li>While still on the dorsal face of the urogenital region, find the base of the seminal vesicle. Carefully remove the seminal vesicle and discard. Perform the same procedure on the opposite side. 
		NOTE: Avoid puncturing the seminal vesicle, as that will release sticky and opaque secretory fluid which interferes with prostate dissection. If the seminal vesicle is punctured, transfer the remaining urogenital region to new 10 cm dish with fresh PBS.</li>
		<li>Remove and discard the vas deferens and as much fatty and connective tissue as possible using the curved side of the forceps.</li>
		<li>While still firmly holding the urethra/proximal prostate region, use a pair of fine, pointed scissors to remove the bladder from the urethra.</li>
	</ol>
	</li>
	<li>Isolation of individual prostate lobes
	<ol>
		<li>Locate the anterior prostate (Figure 1B). Ensure to firmly hold the proximal prostate region/urethra with the straight forceps. Remove the anterior prostate region by directly grasping the tissue with the curved side of the forceps and firmly pulling it away from the bladder and the rest of the prostate. Place the tissue in PBS. 
		NOTE: For all prostate regions, dissecting scissors may be required to remove the tissue, depending on the size of the tumor.</li>
		<li>Locate the ventral prostate region (Figure 1B). Remove the ventral prostate region in the same manner as in step 1.5.7.</li>
		<li>At this point in the dissection, only the lateral and dorsal prostate regions should be present, while the proximal prostate region is still being grasped by the straight forceps. Assess the lateral and dorsal prostate regions (Figure 1B). If the lateral region can be distinguished from the dorsal region, remove the lateral prostate region as described in step 1.5.7. Do the same procedure on the opposite side.</li>
		<li>Remove the dorsal prostate region as described in step 1.5.7. Place the proximal prostate region in 4% PFA, as its structure within the muscular tissue of the urethra makes it unsuitable for organoid generation.</li>
	</ol>
	</li>
</ol>
2. Generation of 3D Organoids from Prostate Tumor Tissue
NOTE: Figure 3 shows a pictorial description of the procedure for generation of tumor organoids.
<ol>
	<li>Preparation
	<ol>
		<li>Prepare mouse prostate organoid media according to Drost et al.18 with the following alterations. Use conditioned medium instead of recombinant proteins for Noggin and R-Spondin and use a final concentration of 1% (v/v), instead of 10%, for both Noggin- and R-Spondin-conditioned medium. 
		NOTE: HEK293 cells stably transfected with HA-mouse Noggin-Fc or HA-mouse Rspo1-Fc are used to produce Noggin- or R-Spondin- conditioned medium, respectively. These cell lines were a gift from the Calvin Kuo Laboratory at Stanford University.</li>
		<li>Prepare digestion solution in a 15 mL tube by diluting 20 mg/mL collagenase II with Advanced DMEM/F12(+++) media to a final concentration of 5 mg/mL. Add Y-27632 Rock Inhibitor to the collagenase II solution at a final concentration of 10 &#181;M. 
		NOTE: The ratio of digestion buffer to tissue is 1 mL to 50 mg, which we use according to Drost et al.18.</li>
	</ol>
	</li>
	<li>Mincing and digestion of tumor tissue
	<ol>
		<li>In the cell culture hood, place prostate tumor tissue in a sterile 10 cm culture dish, dissect and discard necrotic tissue.</li>
		<li>Mince the remaining prostate tumor tissue into 1 mm3 cubes by holding the tissue pieces with the sterile curved forceps and cutting with dissection scissors.</li>
		<li>Place the minced tumor pieces in the 15 mL tube with the digestion buffer by scooping them up with the curved side of the forceps. Digest the tumor tissue at 37 &#176;C with shaking for 1.5 to 2 h. Check digestion progress every 20 min. 
		NOTE: At this time, take out at least 2 mL of matrix from -20 &#176;C storage and thaw on ice. 1 mL aliquots of matrix will take approximately 3 h to thaw.</li>
		<li>After tissue digestion, centrifuge tube at 175 x g for 5 min at 4 &#176;C to form a cell pellet.</li>
		<li>Remove the supernatant, flick the tube to loosen the cell pellet, and resuspend the cell pellet in 1 mL of pre-warmed trypsin supplemented with 10 &#181;M Y-27632 Rock Inhibitor. Put the tube in a 37 &#176;C water bath for 5 min.</li>
		<li>After incubation, pipette up and down 5 times with a standard P1000 tip. Return the tube to 37 &#176;C water bath for another 5 min and repeat step 2.2.6. 
		NOTE: At this time in the procedure, warm a sterile 6-well cell culture dish by putting it into a 55 &#176;C incubator.&#160;</li>
	</ol>
	</li>
	<li>Counting cells and resuspension in matrix
	<ol>
		<li>Wash the cells by adding 9 mL of cold AdDMEM/F12(+++) and centrifuge the tube at 175 x g for 5 min at 4 &#176;C.</li>
		<li>After centrifugation, remove the supernatant, flick the tube to loosen the pellet, and wash the cells again by adding 10 mL of cold AdDMEM/F12(+++). Centrifuge the tube at 175 x g for 5 min at 4 &#176;C.</li>
		<li>After centrifugation, remove the supernatant, flick the tube to loosen the pellet, resuspend the cells in 1 mL of AdDMEM/F12(+++), and count the number of cells using a hemocytometer according to standard procedure.</li>
		<li>Seven-to-eight domes fit in one well of a 6 well dish when organoids are plated in matrix via a drop-wise fashion. Approximately 200 &#956;L of matrix will produce 7-8 domes. Decide how many wells of organoids are needed for future experimental purposes and calculate the volume of matrix required. Then, calculate the volume of cell-containing solution needed to have a final concentration of 1.0 x 106 cells/mL of matrix.</li>
		<li>After counting cells, centrifuge at 175 x g for 5 min at 4 &#176;C. Remove the supernatant, flick the tube to loosen the pellet, and resuspend the cells in volume of matrix calculated in step 2.3.4. 
		NOTE: Matrix remains in liquid form only at 4 &#176;C; keep matrix stock tubes and matrix-cell solutions on ice at all times.</li>
	</ol>
	</li>
	<li>Plating matrix domes and application of media
	<ol>
		<li>Mix matrix-cell solution with a P200 pipet to evenly distribute the cells without introducing bubbles. Remove the 6-well culture dish from the 55 &#176;C incubator.</li>
		<li>Carefully pipette 200 &#956;L of matrix-cell solution and quickly drop the solution into a well to create domes.</li>
		<li>Repeat step 2.4.2. until volume of matrix-cell solution is spent. Allow the domes to solidify at room temperature for 2 min.</li>
		<li>Flip the 6-well dish upside-down and put the dish into a 37 &#176;C incubator to continue solidification for 20 min.</li>
		<li>After incubation, add 2 mL of mouse prostate organoid media to each well. Add synthetic androgen R1881 to each well for a final concentration of 1 nM and Y-27632 Rock Inhibitor to a final concentration of 10 &#956;M. Mix carefully and place the plate in a 37 &#176;C incubator for culturing. 
		NOTE: Organoid culture media needs to be supplemented with 10 &#181;M Y-27632 Rock Inhibitor for only 1 week after organoid generation.</li>
	</ol>
	</li>
</ol>

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The authors do not have financial relationships to disclose.

Critical steps within the protocol for prostate tumor dissection and organoid generation 
Removal of non-prostate tissue and fine dissection of the mouse prostate tumor is crucial for the optimal generation of cancer organoids since both non-prostate epithelial cells and normal prostate epithelial cells will generate organoids. For solid prostate tumors specifically, it is crucial to isolate areas of viable tumor to remove contamination with necrotic tissue that would reduce the number of viable cel...

Since the late 1980s, the ability to alter genes by homologous recombination has greatly advanced the study of biological systems1. Inducible, tissue-, or cell-specific promotor systems and site-specific recombinases, such as Cre-loxP, has advanced genetic studies by facilitating control over genetic modifications both temporally and spatially2,3,4. The combination of these genetic strategies has created a wide array of experimental model systems5,6,7.
Genetically engineered mouse models (GEMMs) are an integral tool to assess how individual genes or groups of genes affect mammalian development and disease. In pre-clinical cancer research, GEMMs are the most physiologically-relevant and rigorous method to study cancer development, progression, and treatment8. Our laboratory specializes in generating and characterizing cancer GEMMs.
The most highly diagnosed non-cutaneous cancer among men in the United States is prostate cancer (PCa). The majority of patients with PCa have low-risk disease and high likelihood of survival, but survival rates decline drastically when disease is diagnosed at advanced stages or if targeted hormonal therapy induces progression to aggressive, non-curable PCa subtypes9,10. Our laboratory has developed GEMMs that utilize floxed alleles of one or more tumor suppressor genes. Recombination and loss of tumor suppressor gene expression occurs specifically in the prostate because we have introduced a transgene with Cre recombinase downstream of the probasin promoter activated only in prostate epithelial cells11,12. We have also bred our GEMMs to contain a Cre reporter transgene called mT/mG, which induces Tomato fluorescent protein expression in cells lacking Cre and green fluorescent protein (GFP) expression in cells with Cre13. While the presentation of this method and our representative results show GEMMs we study in our laboratory, this protocol can be used to generate prostate cancer organoids from any mouse model. However, as discussed in detail in our representative results section, we have observed that certain tumor characteristics are optimal for prostate cancer organoid generation.
In the last decade, new methods of culturing cells from tissues of epithelial origin has led to significant advances in our ability to model organ systems in vitro14,15. The term &#34;3D cell culture&#34;&#160;has been attributed to the techniques involved in establishing and maintaining organoids, which can be generally defined as structures made up of cells that assemble secondary architecture driven by organ-specific cell lineage characteristics16. These new methods are distinct from classic 2D cell culture in that cells do not require transformation or immortalization for long term growth; thus, 3D cultures of normal cells can be compared to diseased cells. This is particularly valuable in cancer research where normal cell control cultures have typically not been available. In addition, organoids spontaneously form secondary tissue architectures with appropriately differentiated cell types, making them a better model system to understand cancer in vitro than 2D cell lines17. Our laboratory has created 3D organoid lines from tumor issue isolated from our PCa GEMMs to complement our in vivo data and perform experiments which would not be feasible in GEMMs.&#160;
In this article, we present written and visual protocols for the complete necropsy of PCa GEMMs, including dissection of distinct mouse prostate lobes and metastatic lesions. We describe and show a step-by-step method for generating organoids from mouse prostate tumors based on a protocol previously published by Drost et al. for deriving organoids from normal mouse prostate epithelial tissue18.

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			<td height="42" style="height:42px;width:217px;">1 1/4 in, 23 gauge, disposable syringe needles</td>
			<td>Becton Dickinson</td>
			<td style="width:119px;">Z192430</td>
			<td></td>
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			<td height="147" style="height:147px;width:217px;">EHS Sarcoma matrix, Pathclear Lot#19814A10</td>
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			<td height="42" style="height:42px;width:217px;">human recombinant Epidermal growth factor (EGF)</td>
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			<td height="21" style="height:21px;width:217px;">Precision balance</td>
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			<td height="42" style="height:42px;width:217px;">Stainless steel dissecting scissors, 10 cm, straight</td>
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			<td height="42" style="height:42px;width:217px;">Stainless steel Iris forceps, 10 cm, curved tip, serrated</td>
			<td style="width:177px;">World Precision Instruments</td>
			<td align="right" style="width:119px;">15915</td>
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			<td height="42" style="height:42px;width:217px;">Stainless steel Nugent utility forceps, straight tip, serrated</td>
			<td style="width:177px;">World Precision Instruments</td>
			<td align="right" style="width:119px;">504489</td>
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			<td height="21" style="height:21px;width:217px;">Y-276632 (Rock Inhibitor)</td>
			<td style="width:177px;">APExBIO</td>
			<td style="width:119px;">A3008</td>
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generation of tumor organoids from genetically engineered mouse models of prostate cancer

Methods based on homologous recombination to modify genes have significantly furthered biological research. Genetically engineered mouse models (GEMMs) are a rigorous method for studying mammalian development and disease. Our laboratory has developed several GEMMs of prostate cancer (PCa) that lack expression of one or multiple tumor suppressor genes using the site-specific Cre-loxP recombinase system and a prostate-specific promoter. In this article, we describe our method for necropsy of these PCa GEMMs, primarily focusing on dissection of mouse prostate tumors. New methods developed over the last decade have facilitated the culture of epithelial-derived cells to model organ systems in vitro in three dimensions. We also detail a 3D cell culture method to generate tumor organoids from mouse PCa GEMMs. Pre-clinical cancer research has been dominated by 2D cell culture and cell line-derived or patient-derived xenograft models. These methods lack tumor microenvironment, a limitation of using these techniques in pre-clinical studies. GEMMs are more physiologically-relevant for understanding tumorigenesis and cancer progression. Tumor organoid culture is an in vitro model system that recapitulates tumor architecture and cell lineage characteristics. In addition, 3D cell culture methods allow for growth of normal cells for comparison to tumor cell cultures, rarely possible using 2D cell culture techniques. In combination, use of GEMMs and 3D cell culture in pre-clinical studies has the potential to improve our understanding of cancer biology.&#160;

Representative necropsy images of a mouse with a large fluid-filled primary prostate tumor in the anterior prostate region are shown in Figure 2A. In contrast, Figure 2B, shows representative necropsy images of a mouse with a large solid primary prostate tumor for which individual prostate regions are indistinguishable. Fluorescent dissection images show the same solid prostate tumor from Figure 2<s...

Watch this Scientific Journal Video about Generation of Tumor Organoids from Genetically Engineered Mouse Models of Prostate Cancer at JoVE.com

Generation of Tumor Organoids from Genetically Engineered Mouse Models of Prostate Cancer

We show a method for necropsy and dissection of mouse prostate cancer models, focusing on prostate tumor dissection. A step-by-step protocol for generation of mouse prostate tumor organoids is also presented.

Methods based on homologous recombination to modify genes have significantly furthered biological research. Genetically engineered mouse models (GEMMs) ...

generation-tumor-organoids-from-genetically-engineered-mouse-models

Research

JoVE Journal

Genetics

10.4K Views.  Roswell Park Comprehensive Cancer Center. We show a method for necropsy and dissection of mouse prostate cancer models, focusing on prostate tumor dissection. A step-by-step protocol for generation of mouse prostate tumor organoids is also presented.

Video: Generation of Tumor Organoids from Genetically Engineered Mouse Models of Prostate Cancer

Generation of Native Chromatin Immunoprecipitation Sequencing Libraries for Nucleosome Density Analysis

We present a modified native chromatin immunoprecipitation sequencing (ChIP-seq) experimental protocol compatible with a Gaussian mixture distribution based analysis methodology (nucleosome density ChIP-seq; ndChIP-seq) that enables the generation of combined measurements of micrococcal nuclease (MNase) accessibility with histone modification genome-wide. Nucleosome position and local density, and the posttranslational modification of their histone subunits, act in concert to regulate local transcription states. Combinatorial measurements of nucleosome accessibility with histone modification generated by ndChIP-seq allows for the simultaneous interrogation of these features. The ndChIP-seq methodology is applicable to small numbers of primary cells inaccessible to cross-linking based ChIP-seq protocols. Taken together, ndChIP-seq enables the measurement of histone modification in combination with local nucleosome density to obtain new insights into shared mechanisms that regulate RNA transcription within rare primary cell populations.

We present a modified native chromatin immunoprecipitation sequencing (ChIP-seq) methodology for the generation of sequence datasets suitable for a nucleosome density ChIP-seq analytical framework integrating micrococcal nuclease (MNase) accessibility with histone modification measurements.

We present a modified native chromatin immunoprecipitation sequencing (ChIP-seq) experimental protocol compatible with a Gaussian mixture distribution ...

Generation of Genome-wide Chromatin Conformation Capture Libraries from Tightly Staged Early Drosophila Embryos

Investigating the three-dimensional architecture of chromatin offers invaluable insight into the mechanisms of gene regulation. Here, we describe a protocol for performing the chromatin conformation capture technique in situ Hi-C on staged Drosophila melanogaster embryo populations. The result is a sequencing library that allows the mapping of all chromatin interactions that occur in the nucleus in a single experiment. Embryo sorting is done manually using a fluorescent stereo microscope and a transgenic fly line containing a nuclear marker. Using this technique, embryo populations from each nuclear division cycle, and with defined cell cycle status, can be obtained with very high purity. The protocol may also be adapted to sort older embryos beyond gastrulation. Sorted embryos are used as inputs for in situ Hi-C. All experiments, including sequencing library preparation, can be completed in five days. The protocol has low input requirements and works reliably using 20 blastoderm stage embryos as input material. The end result is a sequencing library for next generation sequencing. After sequencing, the data can be processed into genome-wide chromatin interaction maps that can be analyzed using a wide range of available tools to gain information about topologically associating domain (TAD) structure, chromatin loops, and chromatin compartments during Drosophila development.

Generation of Genome-wide Chromatin Conformation Capture Libraries from Tightly Staged Early Drosophila Embryos

This work describes a protocol for the generation of high resolution in situ Hi-C libraries from tightly staged pre-gastrulation Drosophila melanogaster embryos.

Investigating the three-dimensional architecture of chromatin offers invaluable insight into the mechanisms of gene regulation. Here, we describe a ...

Simultaneous Video-EEG-ECG Monitoring to Identify Neurocardiac Dysfunction in Mouse Models of Epilepsy

In epilepsy, seizures can evoke cardiac rhythm disturbances such as heart rate changes, conduction blocks, asystoles, and arrhythmias, which can potentially increase risk of sudden unexpected death in epilepsy (SUDEP). Electroencephalography (EEG) and electrocardiography (ECG) are widely used clinical diagnostic tools to monitor for abnormal brain and cardiac rhythms in patients. Here, a technique to simultaneously record video, EEG, and ECG in mice to measure behavior, brain, and cardiac activities, respectively, is described. The technique described herein utilizes a tethered (i.e., wired) recording configuration in which the implanted electrode on the head of the mouse is hard-wired to the recording equipment. Compared to wireless telemetry recording systems, the tethered arrangement possesses several technical advantages such as a greater possible number of channels for recording EEG or other biopotentials; lower electrode costs; and greater frequency bandwidth (i.e., sampling rate) of recordings. The basics of this technique can also be easily modified to accommodate recording other biosignals, such as electromyography (EMG) or plethysmography for assessment of muscle and respiratory activity, respectively. In addition to describing how to perform the EEG-ECG recordings, we also detail methods to quantify the resulting data for seizures, EEG spectral power, cardiac function, and heart rate variability, which we demonstrate in an example experiment using a mouse with epilepsy due to Kcna1 gene deletion. Video-EEG-ECG monitoring in mouse models of epilepsy or other neurological disease provides a powerful tool to identify dysfunction at the level of the brain, heart, or brain-heart interactions.

Here, we present a protocol to record brain and heart bio signals in mice using simultaneous video, electroencephalography (EEG), and electrocardiography (ECG). We also describe methods to analyze the resulting EEG-ECG recordings for seizures, EEG spectral power, cardiac function, and heart rate variability.

In epilepsy, seizures can evoke cardiac rhythm disturbances such as heart rate changes, conduction blocks, asystoles, and arrhythmias, which can ...

Identification of Homologous Recombination Events in Mouse Embryonic Stem Cells Using Southern Blotting and Polymerase Chain Reaction

Relative to the issues of off-target effects and the difficulty of inserting a long DNA fragment in the application of designer nucleases for genome editing, embryonic stem (ES) cell-based gene-targeting technology does not have these shortcomings and is widely used to modify animal/mouse genome ranging from large deletions/insertions to single nucleotide substitutions. Notably, identifying the relatively few homologous recombination (HR) events necessary to obtain desired ES clones is a key step, which demands accurate and reliable methods. Southern blotting and/or conventional PCR are often utilized for this purpose. Here, we describe the detailed procedures of using those two methods to identify HR events that occurred in mouse ES cells in which the endogenous Myh9 gene is intended to be disrupted and replaced by cDNAs encoding other nonmuscle myosin heavy chain IIs (NMHC IIs). The whole procedure of Southern blotting includes the construction of targeting vector(s), electroporation, drug selection, the expansion and storage of ES cells/clones, the preparation, digestion, and blotting of genomic DNA (gDNA), the hybridization and washing of probe(s), and a final step of autoradiography on the X-ray films. PCR can be performed directly with prepared and diluted gDNA. To obtain ideal results, the probes and restriction enzyme (RE) cutting sites for Southern blotting and the primers for PCR should be carefully planned. Though the execution of Southern blotting is time-consuming and labor-intensive and PCR results have false positives, the correct identification by Southern blotting and the rapid screening by PCR allow the sole or combined application of these methods described in this paper to be widely used and consulted by most labs in the identification of genotypes of ES cells and genetically modified animals.

Here, we present a detailed protocol for identifying homologous recombination events that occurred in mouse embryonic stem cells using Southern blotting and/or PCR. This method is exemplified by the generation of nonmuscle myosin II genetic replacement mouse models using traditional embryonic stem cell-based homologous recombination-mediated targeting technology.

Relative to the issues of off-target effects and the difficulty of inserting a long DNA fragment in the application of designer nucleases for genome ...

Generation of Cancer Cell Clones to Visualize Telomeric Repeat-containing RNA TERRA Expressed from a Single Telomere in Living Cells

Telomeres are transcribed, giving rise to telomeric repeat-containing long noncoding RNAs (TERRA), which have been proposed to play important roles in telomere biology, including heterochromatin formation and telomere length homeostasis. Recent findings revealed that TERRA molecules also interact with internal chromosomal regions to regulate gene expression in mouse embryonic stem (ES) cells. In line with this evidence, RNA fluorescence in situ hybridization (RNA-FISH) analyses have shown that only a subset of TERRA transcripts localize at chromosome ends. A better understanding of the dynamics of TERRA molecules will help define their function and mechanisms of action. Here, we describe a method to label and visualize single-telomere TERRA transcripts in cancer cells using the MS2-GFP system. To this aim, we present a protocol to generate stable clones, using the AGS human stomach cancer cell line, containing MS2 sequences integrated at a single subtelomere. Transcription of TERRA from the MS2-tagged telomere results in the expression of MS2-tagged TERRA molecules that are visualized by live-cell fluorescence microscopy upon co-expression of a MS2 RNA-binding protein fused to GFP (MS2-GFP). This approach enables researchers to study the dynamics of single-telomere TERRA molecules in cancer cells, and it can be applied to other cell lines.

Here, we present a protocol to generate cancer cell clones containing a MS2 sequence tag at a single subtelomere. This approach, relying on the MS2-GFP system, enables visualization of the endogenous transcripts of telomeric repeat-containing RNA (TERRA) expressed from a single telomere in living cells.

Telomeres are transcribed, giving rise to telomeric repeat-containing long noncoding RNAs (TERRA), which have been proposed to play important roles in ...

Intraventricular Transplantation of Engineered Neuronal Precursors for In Vivo Neuroarchitecture Studies

Gene control of neuronal cytoarchitecture is currently the subject of intensive investigation. Described here is a simple method developed to study in vivo gene control of neocortical projection neuron morphology. This method is based on (1) in vitro lentiviral engineering of neuronal precursors as "test" and "control" cells, (2) their co-transplantation into wild-type brains, and (3) paired morphometric evaluation of their neuronal derivatives. Specifically, E12.5 pallial precursors from panneuronal, genetically labeled donors, are employed for this purpose. They are engineered to take advantage of selected promoters and tetON/OFF technology, and they are free-hand transplanted into neonatal lateral ventricles. Later, upon immunofluorescence profiling of recipient brains, silhouettes of transplanted neurons are fed into NeurphologyJ open source software, their morphometric parameters are extracted, and average length and branching index are calculated. Compared to other methods, this one offers three main advantages: it permits achieving of fine control of transgene expression at affordable costs, it only requires basic surgical skills, and it provides statistically reliable results upon analysis of a limited number of animals. Because of its design, however, it is not adequate to address non cell-autonomous control of neuroarchitecture. Moreover, it should be preferably used to investigate neurite morphology control after completion of neuronal migration. In its present formulation, this method is exquisitely tuned to investigate gene control of glutamatergic neocortical neuron architecture. Taking advantage of transgenic lines expressing EGFP in other specific neural cell types, it can be re-purposed to address gene control of their architecture.

Based on in vitro lentiviral engineering of neuronal precursors, their co-transplantation into wild-type brains and paired morphometric evaluation of "test" and "control" derivatives, this method allows accurate modeling of in vivo gene control of neocortical neuron morphology in a simple and affordable way.

Gene control of neuronal cytoarchitecture is currently the subject of intensive investigation. Described here is a simple method developed to study in ...

The Use of Mouse Mammary Tumor Cells in an In Vitro Invasion Assay as a Measure of Oncogenic Cell Behavior

The in vitro invasion assay uses a protein-rich matrix in a Boyden chamber to measure the ability of cultured cells to pass through the matrix and a porous membrane in a process analogous to the initial steps of cancer cell metastasis. The tested cells can be altered for the gene expression or treated with inhibitors to test for changes in the invasion potential. This experiment tests the aggressive phenotype of the mouse mammary tumor cells to discover and characterize the potential oncogenes that promote cell invasion. This technique, however, can be versatile and adapted to many different applications. The experiment itself can be done in one day and the results are acquired by light microscopy in less than a day. The results include counts of the number of invading cells for comparison and analysis. The in vitro invasion assay is a rapid, inexpensive, and clear-cut method for determining cell behavior in a culture that can be used as an initial assessment before more involved in vivo assays.

The in vitro cell invasion assay is used to measure the potential of cancer metastasis by quantifying the cellular potential for invasion and migration using cell culture inserts containing protein matrix. Cells are challenged to migrate through the protein matrix and a porous membrane, towards a chemoattractant, and then quantified by light microscopy.

The in vitro invasion assay uses a protein-rich matrix in a Boyden chamber to measure the ability of cultured cells to pass through the matrix and a ...

RNA Next-Generation Sequencing and a Bioinformatics Pipeline to Identify Expressed LINE-1s at the Locus-Specific Level

Long INterspersed Elements-1 (LINEs/L1s) are repetitive elements that can copy and randomly insert in the genome resulting in genomic instability and mutagenesis. Understanding the expression patterns of L1 loci at the individual level will lend to the understanding of the biology of this mutagenic element. This autonomous element makes up a significant portion of the human genome with over 500,000 copies, though 99% are truncated and defective. However, their abundance and dominant number of defective copies make it challenging to identify authentically expressed L1s from L1-related sequences expressed as part of other genes. It is also challenging to identify which specific L1 locus is expressed due to the repetitive nature of the elements. Overcoming these challenges, we present an RNA-Seq bioinformatic approach to identify L1 expression at the locus specific level. In summary, we collect cytoplasmic RNA, select for polyadenylated transcripts, and utilize strand-specific RNA-Seq analyses to uniquely map reads to L1 loci in the human reference genome. We visually curate each L1 locus with uniquely mapped reads to confirm transcription from its own promoter and adjust mapped transcript reads to account for mappability of each individual L1 locus. This approach was applied to a prostate tumor cell line, DU145, to demonstrate the ability of this protocol to detect expression from a small number of the full-length L1 elements.

Here, we present a bioinformatic approach and analyses to identify LINE-1 expression at the locus specific level.

Long INterspersed Elements-1 (LINEs/L1s) are repetitive elements that can copy and randomly insert in the genome resulting in genomic instability and ...

Tumorsphere Derivation and Treatment from Primary Tumor Cells Isolated from Mouse Rhabdomyosarcomas

Rhabdomyosarcoma (RMS) is the most common soft tissue sarcoma in children. Although significant efforts have enabled the identification of common mutations associated with RMS and allowed discrimination of different RMS subtypes, major challenges still exist for the development of novel treatments to further improve prognosis. Although identified by the expression of myogenic markers, there is still significant controversy over whether RMS has myogenic or non-myogenic origins, as the cell of origin is still poorly understood. In the present study, a reliable method is provided for the tumorsphere assay for mouse RMS. The assay is based on functional properties of tumor cells and allows the identification of rare populations in the tumor with tumorigenic functions. Also described are procedures for testing recombinant proteins, integrating transfection protocols with the tumorsphere assay, and evaluating candidate genes involved in tumor development and growth. Described further is a procedure for allograft transplantation of tumorspheres into recipient mice to validate tumorigenic function in vivo. Overall, the described method allows reliable identification and testing of rare RMS tumorigenic populations that can be applied to RMS arising in different contexts. Finally, the protocol can be utilized as a platform for drug screening and future development of therapeutics.

This protocol describes a reproducible method for isolation of mouse rhabdomyosarcoma primary cells, tumorsphere formation and treatment, and allograft transplantation starting from tumorspheres cultures.

Rhabdomyosarcoma (RMS) is the most common soft tissue sarcoma in children. Although significant efforts have enabled the identification of common ...

Use of Freeze-thawed Embryos for High-efficiency Production of Genetically Modified Mice

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. The CRISPR/Cas9 system allows researchers to modify the genome with unprecedented efficiency, fidelity, and simplicity. Harnessing this technology, researchers are seeking a rapid, efficient, and easy protocol for generating GM mice. Here we introduce an improved method for cryopreservation of one-cell embryos that leads to a higher developmental rate of the freeze-thawed embryos. By combining it with optimized electroporation conditions, this protocol allows for the generation of knockout and knock-in mice with high efficiency and low mosaic rates within a short time. Furthermore, we show a step-by-step explanation of our optimized protocol, covering CRISPR reagent preparation, in vitro fertilization, cryopreservation and thawing of one-cell embryos, electroporation of CRISPR reagents, mouse generation, and genotyping of the founders. Using this protocol, researchers should be able to prepare GM mice with unparalleled ease, speed, and efficiency.

Here, we present a modified method for cryopreservation of one-cell embryos as well as a protocol that couples the use of freeze-thawed embryos and electroporation for the efficient generation of genetically modified mice.

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. ...